I am sorry my post was not clear enough.
My tomato leaves were vacuum infilitrated with MG115 (a proteasome inhibitor), water, and FB1 (a fungal toxin). Our lab focuses on disease resistance and plant defense responses. The treatments serve as stress. We wanted to test the relative activity of certain enzymes under those stresses. The two enzymes are B-1,3-glucanase and glutamine synthetase.

I first extracted total protein by adding 1ml of 200mM Tris-HCl buffer (0.25mM EDTA, 5mM DTT, and 1mM PMSF) pH 8.0 to 0.5g of frozen tomato leaves and grinded them using a mortar and pestle. I made sure the mortar and pestle were cooled down with liquid nitrogen. The homogenate was centrifuged at 12000g at 4C for 20min.
The concentration of my protein samples was determined using the standard Bradford method.

B-1,3-glucanase
The reaction mixture consisted of 0.4ml of citric-acid phosphate buffer (pH5.6) containing 1mg/ml laminarin and 0.1ml of enzyme solution (this is my total protein samples, keep in mind that after determining their concentrations, I diluted all to the same final concentration). After 15 min incubation at 37C, 0.5ml of the alkaline copper reagent was added and the mixture was heated at 100C for 10 min. After cooling on ice, 0.5ml of the arsenomolybdate reagent was added, followed by 3.0ml of water after the development of the blue color. The absorbance was measured at 660nm.
Then to measure the relative activity, I set the water treatment as (100%) and I measured the relative activity of the other treatments with respect to water.
Ex. if water-treatment absorbance is 0.56 and the FB1 treatment is 0.97, I divided 0.97/0.56*100= 173.21%

Glutamine Synthetase (GS)
We adapted a method for wheat, even though there are publications with tomato GS methods.
GS activity was measured using the
synthetase assay based on the method described by Lea
et al. (1990) and optimized for wheat leaves as follows.
100uL of crude leaf extract (the same protein samples were used as the ones used in glucanase assay above)was added to 380uL of assay
mix which consisted of 100 mM TEA, 80 mM glutamate,
6 mM hydroxylamine HCl, 20 mM MgSO4, 4 mM EDTA
at pH 7.6. The reaction was started by the addition of 20 uLof 0.2 M ATP at pH 7.6. After 10 min of incubation at
30C, the reaction was stopped by the addition of 500 uL of
ferric chloride reagent (0.24 M TCA, 0.1 M ferric chloride,
1.0 M HCl). Samples were then centrifuged at
10,000 g for 5 min and absorbance read at 505 nm.

We also looked at the relative activity of the enzyme for the different treaments and it was done the same way as that in the glucanase assay above.

I am sorry Bob as this is really long but I wanted to make sure that I am very clear on this. Please if you have more questions or not sure about anything, let me know

Thanks, that was very clear - what you actually need to do is very simple - you need to compare your samples to a standard composed of pure forms or known activities of lysate containing the relevant enzyme (or a similar one) that you can set as the relative 100% strength. Using water as a 100% is not ideal for this sort of thing as it should actually have an activity of 0 (presumably). The water should be treated as a blank to set no activity, it would be better to use the lysis buffer as the blank rather than water, as this will affect the absorbance to some extent.

Thanks for your reply. Hmm..for example for my glucanase assay I did have three controls, each had all the components of the assay except for the substrate, Laminarin. Surprisingly, all after substracting their absorbance from that of blank (in my case my blank is my protein extraction buffer mentioned above in my comment, Tris-HCl buffer) had negative absorbancies.

Also I am still confused why would my lysis buffer result in my negative absorbance values? I am guessing by lysis you mean the protein extraction buffer.

Ok. That is a bit of a problem. I was assuming that you weren't using an extraction buffer control (and I did mean extraction buffer when I talked about lysis buffer). In the case that my assumption was true, the negative absorbances would be due to the buffer being less absorbent than water at the read wavelength.

Would it be possible for there to be something in the plants that might inhibit the reaction (proteases perhaps)?

Sorry Bob in my earlier post I said their absorbance was subtracted from that of the blank, I meant to say the blank's absorbance was subtracted from their absorbance.

About the proteases, I use PMSF in my extraction buffer which should inhibit proteases. Could it be dirt like my microplate being dirty which intereferes with the readings? I am so confused

Also, I don't know if this affects but for my protein samples, I have taken the supernatant after the centrifugation step, although I can still see some residue (really tiny green stuff-leaves maybe) in there.

PMSF is a serine protease inhibitor only, there are quite a number of other proteases (cysteine, metallo, aspartic) out there that won't be inhibited by PMSF at all. I recommend using a cocktail tablet such as Complete from Roche or any suitable one from Sigma-Aldrich. Dirt on some wells would affect your reading by not allowing all the light into the well thereby getting a reduced reading at the end. You could just put an empty plate in the reader and see how it comes out at your wavelengths.

Freeze/thaw of any protein is a good way to degrade activity typically. If you need to do these assays, it is best to make sure that all the lysates have been treated in the same manner, this includes the number of times you have freeze/thawed the lysate. Storage at -80 is also recommended.

Green things in the lysate could well be chlorophyll or chloroplasts (or as you say, bits of leaf), these could absorb in the red spectrum (660 nm is red) but shouldn't absorb in the green (505 nm), though discrete particles could scatter the light which might account for a lower reading than in the blank.